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Autocrine/paracrine pattern of superoxide production through NAD(P)H oxidase in coronary arterial myocytes

¹Department of Pharmacology and Toxicology, Medical College of Virginia, Virginia Commonwealth University, Richmond, Virginia; ²Section of Cardiology, Department of Medicine, Division of Biological Sciences and Pritzker School of Medicine, University of Chicago, Chicago, Illinois; ³ Research Center of Experimental Medicine, Guangxi Autonomous Region People's Hospital, Nanning, Guangxi, People's Republic of China

Submitted 14 June 2006 ; accepted in final form 31 August 2006

	ABSTRACT

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The present study tested the hypothesis that membrane-bound NAD(P)H oxidase in coronary arterial myocytes (CAMs) is capable of producing superoxide (O₂^–) toward extracellular space to exert an autocrine- or paracrine-like action in these cells. Using a high-speed wavelength-switching fluorescent microscopic imaging technique, we simultaneously monitored the binding of dihydroethidium-oxidizing product to exogenous salmon testes DNA trapped outside CAMs and to nuclear DNA as indicators of extra- and intracellular O₂^– production. It was found that a muscarinic agonist oxotremorine (OXO; 80 µM) increased O₂^– levels more rapidly outside than inside CAMs. In the presence of superoxide dismutase (500 U/ml) plus catalase (400 U/ml) and NAD(P)H oxidase inhibitor diphenylene iodonium (50 µM) or apocynin (100 µM), these increases in extra- and intracellular O₂^– levels were substantially abolished or attenuated. The O₂^– increase outside CAMs was also confirmed by detecting oxidation of nitro blue tetrazolium and confocal microscopic localization of Matrigel-trapped OxyBURST H₂HFF Green BSA staining around these cells. By electron spin resonance spectrometry, the extracellular accumulation of O₂^– was demonstrated as a superoxide dismutase-sensitive component outside CAMs. Furthermore, RNA interference of NAD(P)H oxidase subunits Nox1 or p47 markedly blocked OXO-induced increases in both extra- and intracellular O₂^– levels, whereas small inhibitory RNA of Nox4 only attenuated intracellular O₂^– accumulation. These results suggest that Nox1 represents a major NAD(P)H oxidase isoform responsible for extracellular O₂^– production. This rapid extracellular production of O₂^– seems to be unique to OXO-induced M₁-receptor activation, since ANG II-induced intra- and extracellular O₂^– increases in parallel. It is concluded that the outward production of O₂^– via NAD(P)H oxidase in CAMs may represent an important producing pattern for its autocrine or paracrine actions.

reactive oxygen species; redox signaling; artery

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It has become clearer and clearer that there are several potential enzymatic sources of O₂^– in the cardiovascular system, such as xanthine oxidase, mitochondrial respiratory chain arachidonic acid metabolizing system, uncoupled NO synthase, and NAD(P)H oxidase (43, 46). There is increasing evidence that NAD(P)H oxidase is a major source of O₂^– in vascular cells (16, 35, 52) and that NAD(P)H oxidase-derived O₂^– serves as an important physiological redox signaling molecule to participate in the regulation of vascular function. In vascular smooth muscle cells (VSMCs), recent studies have reported that there are membrane-bound and intracellular nonmitochondrial NAD(P)H oxidases, which are all capable of producing O₂^– in these cells (5, 16). With various existing techniques, O₂^– has been demonstrated to be accumulated within VSMCs when NAD(P)H oxidase is activated by different agonists such as ANG II (15). This intracellular accumulation of O₂^– led to an assumption that a plasma membrane-bound NAD(P)H oxidase may produce and release O₂^– into cells, which is different from the orientation of phagocyte NAD(P)H oxidases (16). However, the proposed topology of NAD(P)H oxidase subunits indicates that membrane-associated NAD(P)H oxidase should not release O₂^– into cytosol (4, 25). Studies on subcellular localization of vascular NAD(P)H oxidase subunits also demonstrated that O₂^– within VSMCs may not be derived from plasma membrane NAD(P)H oxidase but rather from intracellular compartmental NAD(P)H oxidase (45, 52). More recently, using patch-clamp technique, our group (30) surprisingly recorded an inward current associated with NAD(P)H oxidase in coronary arterial myocytes (CAMs) that was similar to that recorded in phagocytes, indicating that an outward electron flow and O₂^– production occurred in these VSMCs. It seems that a membrane-bound NAD(P)H oxidase in VSMCs could generate O₂^– toward the outside of these cells, and in this way O₂^– may exert regulatory roles as an autocrine or paracrine.

To test this hypothesis, we first modified a high-speed wavelength-switching fluorescent microscopic imaging technique to allow a simultaneous monitoring of extracellular and intracellular O₂^– production in single CAMs. In addition to the intracellular DNA trapping of dihydroethidium (DHE) product, a Matrigel DNA trapping around CAMs was developed to detect O₂^–-oxidized DHE signals, which represents extracellular O₂^– levels. In addition, O₂^–-mediated precipitation from nitro blue tetrazolium (NBT) and Matrigel-trapped OxyBURST H₂HFF Green BSA staining around CAMs were also determined by light or confocal fluorescent microscopy, respectively. Using electron spin resonance (ESR) spectrometric measurement, we examined whether SOD-sensitive extracellular O₂^– changes could be detected. After we confirmed the extracellular production of O₂^– in response to activation of M₁-muscarinic receptor, we observed the effects of O₂^– scavengers or specific NAD(P)H oxidase inhibitors on intracellular and extracellular O₂^– production and also dissected the contribution of different NAD(P)H oxidase isoforms to this agonist-induced redox response by RNA interference with specific, small inhibitory RNAs (siRNAs). We then went on to examine whether O₂^– outward production in response to M₁-muscarinic activation occurs when CAMs were stimulated by other agonists, in particular ANG II. Our results suggest that activation of NAD(P)H oxidase gives rise to a different spatiotemporal pattern of O₂^– production when these cells are exposed to different stimuli. Most importantly, our experiments provide direct evidence at the single cell level that the outward production of O₂^– associated with NAD(P)H oxidase occurs in CAMs, which may exert autocrine or paracrine actions in the redox regulation of vascular function.

	MATERIALS AND METHODS

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Isolation and culture of CAMs. As our group described previously (41), fresh bovine hearts were obtained from a local abattoir, and intramyocardial coronary arteries from the branches of left anterior descending artery were rapidly dissected. Endothelium-denuded coronary arteries were rinsed with medium 199 containing 5% FBS, a 2% solution of antibiotics (penicillin-streptomycin), 0.3% gentamicin, and 0.3% nystatin and cleaned of connective tissues. After they were cut into small pieces, the arteries were placed into dishes with the lumen side down and incubated in medium 199 containing 10% FBS, 1% glutamine, 1% antibiotic solution, 0.3% gentamicin, 0.3% nystatin, and 0.1% tylosin for 3–5 days until vascular myocytes migrated to the dishes. Once CAM growth had been established, the vessels were removed, and the cells were grown in medium 199 containing 20% FBS. All cells were maintained in an incubator with 5% CO₂ in air at 37°C. The CAMs were identified by positive staining with an anti--actin antibody. All studies were performed with cells of passages 2–4.

Simultaneously monitoring O₂^– production inside and outside CAMs. This method is based on the DNA binding characteristic of oxidized DHE product ethidium in intracellular DNA and Matrigel-trapped extracellular DNA when CAMs were stimulated by different agonists. This DNA binding assay of O₂^–, which oxidizes DHE to produce a strong red fluorescence, is widely used for intracellular O₂^– production in the vessel wall and in isolated vascular endothelial and VSMCs (6, 40). However, this DNA-ethidium binding assay has not yet been used to detect extracellular O₂^– at the single cell level. The major challenge is how to trap DNA outside the cells. To solve this problem, we have used Matrigel (BD Biosciences) as a supportive matrix to trap DNA that is capable of binding to the ethidium from oxidation of DHE in solution. In these experiments, isolated CAMs (10² cells/well) were seeded into a 16-well chamber slide with a transparent glass bottom (LAB-TEK) and incubated overnight to attach the bottom of the wells. On the day of experiments, the culture medium was first discarded, CAMs were washed twice with Hanks’ buffer, and 40 µl of salmon testes DNA solution (7.5 mg/ml; Sigma) were mixed with 40 µl of Matrigel solution (at 4°C) and then were carefully loaded onto the top of CAMs. After 5 min at room temperature, the gel was polymerized and exogenous DNA was immobilized around these cells. The CAMs with Matrigel were then filled with Hanks’ buffer containing 250 µM DHE. After a 60-min loading of DHE into the CAMs, the chamber slide was mounted on the stage of a fluorescent microscope that we routinely used for high-speed wavelength-switching imaging acquisition and recording. For both intracellular and extracellular DHE oxidizing signals, a ratio of ethidium-DNA to DHE signals was recorded (ethidium-DNA fluorescence signal was detected at an excitation of 480 nm and an emission of 610 nm, and DHE fluorescence signal was detected at an excitation of 380 nm and an emission of 445 nm). For each experiment, 8–10 CAMs were monitored simultaneously and an average value was used for statistical analysis.

The M₁-receptor agonist oxotremorine (OXO; 80 µM) was added to activate its receptor and consequently stimulate O₂^– production. In the groups of CAMs receiving O₂^– scavengers or NAD(P)H oxidase inhibitors, SOD (500 U/ml) plus catalase (400 U/ml), diphenylene iodonium (DPI; 50 µM), or apocynin (100 µM) was preincubated with the cells for 15 min, and then the O₂^–-producing response to OXO was determined. In addition, a time course of NAD(P)H oxidase activation and intracellular and extracellular O₂^– production in response to ANG II was monitored, when this peptide was added into the well containing Matrigel and cells at a concentration of 100 nM. All doses of these agonists or inhibitors used in the present experiments were based on previous studies reported by our laboratory and other laboratories.

NBT precipitation detection of extracellular O₂^–. As described in previous studies (13), O₂^– accumulation outside CAMs was also determined with NBT, which reacts with O₂^– and produces a blue formazin precipitate. NBT (0.1%) was prepared in PBS by adding 10 mg of NBT powder (Sigma) to 100 ml of PBS (pH 7.2) and stirred at room temperature for 1 h. NBT staining was then performed by adding 0.1% of NBT solution into a well seeded with CAMs and incubated for 30 min at 37°C before and after addition of OXO (80 µM). Under x100 magnification microscopy (Nikon), two-dimensional optical images were acquired with a digital camera (Spot).

Confocal microscopic detection of extracellular O₂^–. To further determine extracellular O₂^–, CAMs were prepared as described above for ethidium-DNA trapping imaging. In brief, 0.5 ml of Matrigel was mixed with 25 µl of OxyBURST H₂HFF Green BSA (10 µg/ml; Molecular Probes) and then used to evenly coat the 35 mm x 10-mm cell culture dish. Freshly isolated CAM suspension (60 µl) was loaded on the top of the ice-cold Matrigel matrix, and then the culture dish was gently tapped three or four times to sit cells on the surface of the gel. At room temperature, the gel was polymerized in 5 min. Then, 2 ml of Krebs-Ringer phosphate buffer, which contains PBS, pH 7.4, with 1.0 mM Ca²⁺, 1.5 mM Mg²⁺, and 5.5 mM glucose, were added. Confocal fluorescent microscopic images were acquired by an Olympus Fluoview system (version 4.2, FV300), consisting of an Olympus BX61WI inverted microscope with an Olympus Lumplan F1 x60, 0.9 numerical aperture, water-immersion objective. A single z-section was taken, or 0.1-µm sections were obtained through the cell with excitation and emission wavelengths of 488 and 530 nm, respectively, for OxyBURST. Real-time microscopic fluorescence images were acquired every 1 or 2 min under control conditions and after different treatments. This OxyBURST H₂HFF Green BSA detects H₂O₂, which mirrors O₂^– production outside the cells because BSA cannot enter the cells (8).

ESR detection of extracellular O₂^–. For extracellular O₂^– assay, gently collected CAMs were suspended in modified Krebs-HEPES buffer containing deferoxamine (100 µM; metal chelator). Approximately 1 x 10⁶ cells were then incubated with OXO (80 µM) or ANG II (100 nM) for 30 min; these were subsequently mixed with 1 mM of the O₂^–-specific spin trap 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine (CMH) in the presence or absence of manganese-dependent SOD (500 U/ml). The cell mixture was then loaded in glass capillaries and immediately kinetically analyzed for O₂^– production for 10 min. The SOD-inhibitable fraction of the signal was compared. The ESR settings were as follows: biofield, 3,350; field sweep, 60 G; microwave frequency, 9.78 GHz; microwave power, 20 mW; modulation amplitude, 3 G; 4,096 points of resolution; receiver gain, 100; and kinetic time, 10 min (7).

RNA interference and Nox1, Nox4, and p47^phox gene in CAMs. To dissect the role of different NAD(P)H oxidase subunits in OXO-induced or ANG II-induced O₂^– production outside or inside CAMs, RNA interference was performed to silence the gene coding these subunits. Three pairs of siRNAs were designed and targeted, including p47 (AAAGGGCTCGAGTTCCCAAAT), Nox1 (AAGGGCTTTCGAACAACAATA), Nox4 (AAGACCTGGCCAGTATATTAT), and Nox2 (AAGTCGGTCTGGTACAAATAT). Selection of these NAD(P)H oxidase subunits was based on previous studies that indicated that these may contribute to the activity of membrane-bound and cytosolic compartmental components of NAD(P)H oxidases (20, 45). In these experiments, siRNA transfection was performed according to the manufacturer's instruction in Qiagen TransMessenger kit. All siRNAs were chosen with a Qiagen siRNA design program and synthesized and double-stranded by Xeragon. A scrambled, small RNA was synthesized for a negative control.

Statistics. Data are expressed as means ± SE. The significance of the differences in mean values between and within multiple groups such as multiple time points was examined by ANOVA for repeated measures followed by Duncan's multiple range test. Student's t-test was used to evaluate the statistical significance of differences between two paired observations. P < 0.05 was considered statistically significant.

	RESULTS

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Spatiotemporal pattern of O₂^– production by CAMs in response to OXO. Using the high-speed wavelength-switching technique, we simultaneously monitored the fluorescence intensity of ethidium-DNA complex within primary cultures of CAMs and in Matrigel around these cells. Figure 1A shows typical fluorescent images acquired during 30 min when a CAM received OXO at a concentration of 80 µM. It was found that, at the first 10 min of incubation with OXO, O₂^– levels increased with a predominate response outside cells (becoming red). Starting at 15 min of OXO incubation, intracellular fluorescence intensity increased when the outside signal became much stronger. At 30 min of OXO incubation, intracellular fluorescence became as intense as those observed outside cells. This dynamic change in intracellular and extracellular ethidium-DNA fluorescence was digitized and acquired by using a MetaFluor program (version 6.2). Summarized results are presented in Fig. 1B. It was clear that OXO produced a more rapid increase in ethidium-DNA fluorescence outside CAMs than inside these cells. At 30 min of OXO incubation, the intracellular signal reached a level similar to that observed outside CAMs. In addition, we also monitored similar changes in dynamic responses of intracellular and extracellular ethidium-DNA fluorescence in freshly isolated CAMs in response to OXO (data not shown).

View larger version (30K): [in this window] [in a new window]   Fig. 1. Fluorescent microscopic imaging analysis to simultaneously monitor oxotremorine (OXO)-induced superoxide (O2–) production inside and outside single coronary arterial myocytes (CAMs). A: typical time-dependent increase in oxidized dihydroethidium (DHE) fluorescence ratio [ethidium bromide (EtBr)/DHE] image inside and outside a CAM in the presence or absence of superoxide dismutase (SOD; 500 U/ml). Red fluorescence image indicates O2– level higher than blue background. B: summarized digitized data showing the spatiotemporal pattern of O2– increase outside and inside CAMs during OXO (80 µM) stimulation. C: spatiotemporal pattern of O2– increase outside and inside CAMs during OXO stimulation after treatment with SOD (500 U/ml) in combination with catalase (CAT; 400 U/ml) (n = 6 cases). *P < 0.05 compared with control (at 0 min); #P < 0.05 compared with values obtained outside CAMs.

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To address a concern regarding whether a rapid increase in extracellular fluorescent signal in response to OXO stimulation may be due to the different speeds of ethidium binding to intracellular DNA or extracellular DNA in Matrigel, we performed a similar recording with substitution of intracellular 2',7'-dichlorofluorescin diacetate (H₂DCFDA, a cell-permeant indicator for reactive oxygen species) for DHE, where H₂DCFDA in cytosol directly reacts with H₂O₂. The results showed a similar pattern of intracellular and extracellular changes in O₂^– production in response to OXO (data not shown), where the extracellular increase in fluorescent signal was more rapid than the intracellular increase.

NBT precipitation. As shown in Fig. 2A, using NBT staining, we found that dark blue precipitations were significantly increased when CAMs were incubated with OXO. These experiments were performed to further confirm an extracellular production of O₂^–. As mentioned above, NBT in the bath solution could be reduced by reactive oxygen species (ROS) to dark blue, insoluble formazin-NBT, producing dark blue particles outside the cells. This dye is widely used to detect activation of NAD(P)H oxidase in different cells (12, 18). Although NBT may not be used to specifically dissect which ROS molecules are produced outside CAMs, when it was reduced and precipitated in these experiments, it is indicative for O₂^– production under these conditions.

View larger version (47K): [in this window] [in a new window]   Fig. 2. Microscopic confirmation of extracellular O2– production in CAMs. A: nitro blue tetrazolium staining, where dark blue precipitations were found outside CAMs when they were stimulated by OXO (80 µM). B: confocal fluorescent images showing extracellular O2– production as measured by OxyBURST Green H2HFF BSA trapped H2O2 outside a CAM exposed to OXO (80 µM) in the absence or presence of SOD (500 U/ml) plus CAT (400 U/ml) (n = 5 cases).

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ESR measurements of extracellular O₂^– levels. Using ESR spectrometric assay, we demonstrated the O₂^– production and release from CAMs in response to OXO. In these experiments, a 30-min incubation of OXO with CAMs elicited a typical ESR signal resulting from the CMH trapping, which was measured from cultured CAMs rapidly collected after exposure to OXO. The SOD-inhibitable fraction of the signal, specifically reflecting extracellular O₂^– production, was compared. Figure 3A depicts representative changes in SOD-inhibitable ESR spectrometric curve recorded under control conditions and after treatment of cells with OXO (80 µM) or ANG II (100 nM). Summarized data from these experiments showed that both OXO and ANG II significantly increased production of O₂^– outside CAMs (Fig. 3B). At 10 min, a more than threefold increase in extracellular O₂^– production was observed in these ANG II- or OXO-treated CAMs.

View larger version (17K): [in this window] [in a new window]   Fig. 3. Electron spin resonance (ESR) spectrometric assay of extracellular O2– levels produced by CAMs. A: representative ESR spectrograph of O2– trapped by 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine (CMH) showing SOD-sensitive component of signals recorded under control conditions and after treatment of cells with OXO (80 µM) or ANG II (100 nM). B: summarized data showing ANG II or OXO-induced O2– production (n = 6 cases). *P < 0.05 compared with control.

Effects of NAD(P)H oxidase inhibitors on OXO-induced O₂^– production.

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View larger version (12K): [in this window] [in a new window]   Fig. 4. Effects of NAD(P)H oxidase inhibitor on OXO-induced O2– production inside and outside CAMs as measured by fluorescent microscopic imaging analysis. A: inhibition of OXO (80 µM)-induced O2– production by diphenylene iodonium (DPI; 50 µM) (n = 6 cases). B: inhibition of OXO-induced O2– production by apocynin (APO; 100 µM) (n = 6 cases). *P < 0.05 compared with control (at 0 min).

Effects of gene silencing of NAD(P)H oxidase subunits on OXO-induced O₂^– production.

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View larger version (12K): [in this window] [in a new window]   Fig. 5. Effects of RNA interference on OXO (80 µM)-induced O2– production inside and outside CAMs as measured by fluorescent microscopic imaging analysis. A: effect of Nox1 small inhibitory RNA (siRNA) or scrambled, small RNA (n = 6 cases). B: effect of Nox4 siRNA or scrambled, small RNA (n = 6 cases). C: effect of p47phox siRNA or scrambled, small RNA (n = 6 cases). *P < 0.05 compared with values obtained from CAMs treated with scrambled, small RNA.

Effects of exogenous O₂^– on intracellular NAD(P)H oxidase activity.

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View larger version (9K): [in this window] [in a new window]   Fig. 6. Fluorescent microscopic imaging analysis to simultaneously monitor X/XO-induced O2– production inside and outside single CAMs. A: time-dependent increase in oxidized DHE fluorescence ratio (EtBr/DHE) inside and outside CAMs before and after treatment with X (100 µM)/XO (0.1 mU/ml). B: in the presence of DPI (50 µM), X/XO induced EtBr/DHE signal inside and outside CAMs. C: in the presence of APO (100 µM), X/XO induced EtBr/DHE signal inside and outside CAMs. In A–C, n = 6 cases. *P < 0.05 compared with control (at 0 min); #P < 0.05 compared with values obtained outside CAMs.

Spatiotemporal pattern of ANG II-induced O₂^– production in CAMs.

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View larger version (30K): [in this window] [in a new window]   Fig. 7. Fluorescent microscopic imaging analysis to simultaneously monitor ANG II (100 nM)-induced O2– production inside and outside CAMs. A: typical time-dependent increase in oxidized DHE fluorescence ratio (EtBr/DHE) image inside and outside CAM in the presence or absence of SOD (500 U/ml). Red fluorescence image indicates O2– level higher than blue background. B: summarized digitized data showing the spatiotemporal pattern of O2– increase outside and inside CAMs during ANG II stimulation. C: spatiotemporal pattern of O2– increase outside and inside CAMs during ANG II stimulation after treatment with SOD (500 U/ml) in combination with CAT (400 U/ml) (n = 6 cases). *P < 0.05 compared with control (at 0 min).

Effects of NAD(P)H oxidase inhibitors on ANG II-induced O₂^– production.

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View larger version (12K): [in this window] [in a new window]   Fig. 8. Effects of NAD(P)H oxidase inhibitor on ANG II (100 nM)-induced O2– production inside and outside CAMs as measured by fluorescent microscopic imaging analysis. A: inhibition of ANG II-induced O2– production by DPI (50 µM) (n = 6 cases). B: inhibition of ANG II-induced O2– production by APO (100 µM) (n = 6 cases). *P < 0.05 compared with control (at 0 min).

Effects of gene silencing of NAD(P)H oxidase subunits on ANG II-induced O₂^– production.

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View larger version (15K): [in this window] [in a new window]   Fig. 9. Effects of RNA interference of NAD(P)H oxidase on ANG II-induced O2– production inside and outside CAMs as measured by fluorescent microscopic imaging analysis. A: effect of Nox1 siRNA or scrambled, small RNA (n = 6 cases). B: effect of Nox4 siRNA or scrambled, small RNA (n = 6 cases). C: effect of p47phox siRNA or scrambled, small RNA (n = 6 cases). *P < 0.05 compared with values obtained from CAMs treated with scrambled, small RNA.

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

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It should be noted that the major goal of our study is to determine the role of O₂^– in the rapid and early vasomotor responses to different agonists. Although many activators can stimulate O₂^– production, they are usually used to stimulate vascular O₂^– generation or accumulation for a relatively longer time. On the basis of our previous studies, bovine coronary arteries did not produce vasoconstrictor responses to many agonists such as ANG II, norepinephrine, TNF-, Fas ligand, and other inflammatory cytokines (56, 57). In preliminary experiments, we tried several vasoconstrictors of bovine coronary arteries, including OXO, U-46619, KCl, and CaCl₂, which all caused O₂^– production. Therefore, OXO was chosen as a prototype agonist of bovine coronary vasoconstrictors, which has been often used in previous studies (14, 27, 47). Although ANG II does not strongly constrict bovine coronary arteries, it was used for comparison with OXO in the present study because it has been extensively used as an NAD(P)H oxidase stimulator.

To study the spatiotemporal pattern of O₂^– production associated with NAD(P)H oxidase activity, a sensitive and dynamic measurement of O₂^– both inside and outside cells is needed. By modification of a high-speed wavelength-switching fluorescent microscopic imaging system that we used for simultaneous monitoring of Ca²⁺ and NO in vascular cells (51) and with the help of Matrigel to trap DNA around CAMs, we developed a highly sensitive imaging method to simultaneously monitor extracellular and intracellular DHE-oxidizing signals, which represent O₂^– levels. In these experiments, Matrigel as a supportive matrix trapped DNA that is capable of binding to ethidium from oxidation of DHE in solution, and a ratio of ethidium-DNA to DHE fluorescent signals was recorded to represent O₂^–. This ratiometric assay of O₂^– levels using DHE was used in our previous study and by others (36, 55); this method increases the sensitivity for O₂^– detection and avoids an artifact in assays from cell volume changes or contractions during agonist stimulation. By high-speed wavelength switching, extracellular and intracellular signals can be simultaneously recorded. Using this record system, we first found that, in response to M₁-receptor activation, extracellular and intracellular O₂^– signals could be resolved in a time-dependent manner, which was blocked by the presence of SOD and CAT, suggesting that this assay can be used for dynamic analyses of O₂^– production. In particular, it seems that the system is sensitive for detecting O₂^– production of a single cell at an early stage of stimulation by M₁ agonist. In previous studies, although DHE-based assay was used for a variety of samples or preparations for detection of intracellular O₂^–, a dynamic observation of O₂^– production within a short time period of 10 or 30 min could not be resolved (2). More importantly, simultaneously monitoring of O₂^– increases inside and outside CAMs in response to M₁ agonist provides an important tool to study the orientation of O₂^– production in these cells and to explore the triggering mechanism of redox signaling in the regulation of vascular function. For example, the finding of a relatively early SOD-blockable increase in O₂^– levels outside CAMs may suggest that outward O₂^– production in the early stage of M₁-receptor activation triggers intracellular increases in O₂^– because addition of cell membrane-impermeable SOD outside cells blocked increases in O₂^– signals both outside and inside CAMs. In this regard, O₂^– may serve as an autocrine to activate some membrane receptors or different signaling pathways to result in O₂^– production from intracellular sources (45). In addition, O₂^– can also enter the cells through its channels or other membrane carriers and activate intracellular O₂^– production through different enzymes such as NAD(P)H oxidase. Recent studies have already demonstrated that NAD(P)H oxidase could be activated by O₂^–, leading to a positive feedback response (31). In the present study, we also tested whether extracellular O₂^– stimulates NAD(P)H oxidase activation. It was found that addition of X/XO in the bath solution significantly induced extracellular and intracellular O₂^– production with a pattern similar to that of OXO, which was markedly attenuated by NAD(P)H oxidase inhibitors. This indicates that O₂^– from X/XO outside these cells at least partially exerts its action through enhanced intracellular O₂^– production. This stimulatory action of O₂^– on intracellular NAD(P)H oxidase may explain previous findings from our laboratory and by others that extracellular addition of the O₂^–-producing system largely increased intracellular O₂^– (38, 58). Given the ionic characteristics of O₂^–, the entry of O₂^– into cells should be relatively difficult. Therefore, increased intracellular O₂^– under this condition would be primarily produced from activation of intracellular NAD(P)H oxidase. In addition, this redox activation of NAD(P)H oxidase by another ROS, H₂O₂, has also been reported recently (31).

There was a concern that the temporal difference between increased O₂^– induced by M₁ agonist inside and outside cells may be due to different trapping or trafficking efficiency of intra- or extracellular O₂^–, since the DHE-oxidizing product ethidium needs to bind to DNA. However, some of our results demonstrated that this is not the case. First, we used another indicator, H₂DCFDA, which directly detects O₂^– metabolite via SOD (H₂O₂) in cytoplasma and found a pattern of changes in intracellular and extracellular redox signals similar to what was observed with DHE as the indicator inside and outside CAMs. Second, when another agonist (ANG II) was used, a parallel increase in O₂^– inside and outside these cells was observed. This is different from the pattern exhibited during M₁-receptor activation. These results further confirm that the spatiotemporal patterns of O₂^– production observed by this monitoring system represent different responses of intracellular and extracellular production, rather than the artifact of the assays.

One of the major goals in the present study is to illustrate the production of O₂^– outside CAMs because there is no direct evidence showing that vascular myocytes could produce O₂^– toward the outside of these cells, despite this orientation of O₂^– production being assumed (4, 25). In addition to fluorescent imaging analysis, the present study used three other different methods to demonstrate that CAMs could enzymatically produce O₂^– toward the outside of these cells. These methods included microscopic detection of NBT staining, confocal microscopy analysis of extracellularly trapped H₂O₂ by Matrigel, and ESR measurements. It was found that M₁ agonist OXO induced formation of insoluble formazin-NBT precipitation on the surface of CAMs. This dye was widely used to determine activation of NAD(P)H oxidase in phagocytes for O₂^– production during respiratory burst (39). Using confocal microscopy, we observed an increase in green fluorescence of OxyBURST H₂HFF BSA dye around CAMs in response to OXO, which indicates extracellular H₂O₂ increase (8). Because this BSA conjugate cannot enter cells, this detected green fluorescence represents extracellular ROS. Although this assay did not directly detect O₂^– levels, the increase in H₂O₂ amount outside these cells under this short-term OXO stimulation is undoubtedly indicative of O₂^– production or accumulation, since there is evidence that inhibition of NAD(P)H oxidase could attenuate the enhancement of this extracellular fluorescence derived from the BSA-H₂HFF conjugates. Finally, we performed ESR spectrometry to further confirm OXO-induced O₂^– production outside these CAMs. In these experiments, SOD-sensitive components of ESR spectrometric signals in cell suspensions were analyzed to represent extracellular O₂^–. Given the high specificity of ESR trapping assay in detecting O₂^–, these experiments attempted to confirm that O₂^– is detectable outside CAMS when they are stimulated by agonists. Although this ESR spectrometric assay could not use a single cell to detect O₂^– level (9), it is important that the signal highly specific to O₂^– can be detected outside CAMs, which can further support the view that an outward production of O₂^– occurs in these cells in response to M₁-receptor activation. The results showed that the SOD-sensitive ESR signal was significantly increased when CAMs were stimulated by OXO. Taken together, all four different assays that we used in the present study consistently demonstrated that there is a mechanism by which O₂^– is produced toward the outside of CAMs when their M₁ receptors are activated. This outwardly produced O₂^– may play an autocrine or paracrine role in the regulation of vascular function.

To determine the contribution of NAD(P)H oxidase to increased extra- and intracellular O₂^– production, we first examined the effects of two widely used NAD(P)H oxidase inhibitors (DPI and apocynin) on O₂^– increases induced by OXO. Fluorescent imaging analysis demonstrated that, in the presence of either DPI or apocynin, OXO-induced increases in O₂^– levels both outside and inside CAMs were substantially blocked. In particular, increased intracellular O₂^– under this condition was completely abolished. This suggests that both extra- and intracellular O₂^– are associated with the activity of NAD(P)H oxidase. Next, we examined the effects of RNA interference by specific siRNA of different subunits of NAD(P)H oxidase. It was found that introduction of Nox1 or p47^phox-specific siRNA significantly attenuated OXO-induced increases in O₂^– production both inside and outside CAMs. However, when these cells were transferred with Nox4 siRNA, only the intracellular increase in O₂^– was reduced. This Nox4 RNA interference had no effect on increased production of extracellular O₂^–. These results provide evidence that both extracellular and intracellular O₂^– production are associated with NAD(P)H oxidase activity and that different mechanisms or isoforms of NAD(P)H oxidase may mediate O₂^– production inside or outside these arterial myocytes. It seems that Nox1 rather than Nox4 is primarily responsible for the production of extracellular O₂^–, since Nox4 siRNA and Nox2 could not block the increase in O₂^– outside CAMs. Although the attenuation of intra- and extracellular increases in O₂^– by Nox1 siRNA seems to indicate that this isoform of NAD(P)H oxidase contributes to the production of O₂^– both outside and inside these cells, as discussed above, the increase in intracellular O₂^– via Nox1 may not be directly toward the inside of these cells. It is possible that, in CAMs, Nox1 is activated by stimulation of M₁ receptors and then O₂^–, as an autocrine or paracrine, may lead to activation of an intracellular NAD(P)H oxidase, which may be Nox4 because Nox2 siRNA had no effect on increases in intracellular O₂^–. Several previous reports have indicated that Nox4 is a primary isoform responsible for intracellular O₂^– production (44, 52). In this regard, Nox1 has been shown to be enriched in the membrane fraction and Nox4 predominantly in the intracellular compartments of vascular myocytes (20, 52). It is known that Nox1 is membrane bound and uses intracellular NADPH or NADH as substrate to produce O₂^–. Although there is a possibility that an extracellular NAD(P)H oxidase may be present (37), detectable O₂^– outside CAMs in our preparations is mainly from an enzyme located inside cells because extracellular addition of NADPH (10 mM) was not found to induce any changes in O₂^– production both inside and outside these cells. There are many possible mechanisms by which extracellular O₂^– may further activate intracellular NAD(P)H oxidase to produce intracellular O₂^–. For example, extracellular O₂^– has been reported to activate PKC and thereby result in phosphorylation of p47^phox, thus activating intracellular NAD(P)H (4). In other studies, extracellular O₂^– was found to alter the activity of cell membrane ion channels such as inhibition of K⁺ channels or activation of Ca²⁺ channels and increase intracellular Ca²⁺ concentrations. Because NAD(P)H oxidase activity is Ca²⁺ sensitive, increased Ca²⁺ would activate NAD(P)H oxidase activity (1). In addition, extracellular O₂^– could be converted into H₂O₂, which can easily enter the cells to directly or through some other pathway activate NAD(P)H oxidase, resulting in intracellular O₂^– production (31).

Another important finding of the present study was that ANG II stimulates CAMs to produce O₂^– outside and inside CAMs in parallel, which is different from that observed when these cells were stimulated by OXO. This parallel increase in extracellular and intracellular O₂^– production in response to ANG II could be blocked or substantially attenuated by either SOD plus CAT or NAD(P)H oxidase inhibitors, suggesting that NAD(P)H oxidase is also responsible for the O₂^– production both outside and inside these arterial myocytes during ANG II stimulation. It is unclear why ANG II increases extracellular and intracellular O₂^– production in parallel, whereas OXO increases O₂^– production earlier extracellularly than intracellularly. One possible mechanism may be associated with their postreceptor activating pathways. In this regard, ANG II has been demonstrated to activate NAD(P)H oxidase through phosphorylation of its subunits such as p47^phox (28). Given no specificity of this phosphorylation process to cellular compartments, NAD(P)H oxidase could be activated no matter where this enzyme is primarily located, either on the cell membrane or in intracellular compartments such as the sarcoplasmic reticulum (52), lysosomes (8), and even cytosol (17). Therefore, a parallel increase in extracellular and intracellular O₂^– could be observed when ANG II was added. However, the M₁ agonist OXO may activate NAD(P)H oxidase through changes in cell membrane electrical feature such as depolarization of membrane, rather than phosphorylation. NAD(P)H oxidase electrogenic activation has been reported in some other cells (11, 29). This membrane electrogenic activation may lead to a rapid production of O₂^– toward the outside of cells followed by activation of intracellular NAD(P)H oxidase through the autocrine action of O₂^– or its metabolites outside these cells. Taken together, our results suggest that membrane-bound and intracellular organelle NAD(P)H oxidases could be activated with a temporal difference when arterial myocytes receive different agonistic stimuli.

It should be noted that NAD(P)H oxidase activation may not require p47^phox in some other cell types. For example, Martyn et al. (33) reported that this phox subunit is not needed for NAD(P)H oxidase activation in epithelial cell. We also found that, in renal mesangial cells, p47^phox translocation from cytosol to membrane does not occur during different stimuli (50). However, this p47^phox translocation is strongly required by bovine coronary endothelial cells (59). In VSMCs, considerable evidence shows that p47^phox is needed during activation of NAD(P)H oxidase (24). Our results using p47^phox siRNA demonstrate that both intracellular and extracellular O₂^– productions require p47^phox.

In summary, the present study demonstrated the following. 1) M₁-receptor activation stimulated O₂^– production both outside and inside CAMs with a spatiotemporal pattern of more rapid extracellular increase in O₂^– levels followed by a slow increase in intracellular O₂^– levels. This O₂^– production induced by M₁ agonist is due to NAD(P)H oxidase activation. 2) Nox1 was found to primarily contribute to extracellular O₂^– production, whereas Nox4 is a dominant enzyme responsible for intracellular O₂^– generations. 3) ANG II also stimulated both extra- and intracellular O₂^– production, but the spatiotemporal pattern was different from that observed during M₁ agonist stimulation, which increases O₂^– levels outside and inside CAMs in parallel. This difference of the spatiotemporal pattern of O₂^– production in CAMs in response to different agonists may be related to different activating mechanism of NAD(P)H oxidase when these cells receive stimuli. It is concluded that, in response to different agonists, CAMs exhibit different spatiotemporal patterns of O₂^– production, which is associated with NAD(P)H oxidase activation. Extracellular O₂^– production through Nox1 on the cell membrane represents an autocrine or paracrine model by which O₂^– may exert an important role in autocrine regulation or cell-cell communication in coronary arteries.

	GRANTS

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This study was supported by National Heart, Lung, and Blood Institute Grants HL-57244, HL-75316, and HL-70726.

	FOOTNOTES

 

Address for reprint requests and other correspondence: P.-L. Li, Dept. of Pharmacology and Toxicology, Medical College of Virginia, 410 North 12th St., Richmond, VA 23298 (e-mail: pli{at}vcu.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

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